The human ovarian cancer cell line PEO-14 was obtained from the European Collection of Authenticated Cell Cultures. CAOV3, human ovarian cancer cells, and Chinese hamster ovarian cells (CHO) were purchased from an American Type Culture Collection. Human umbilical vein endothelial cells (HUVEC) were purchased from Lonza. CAOV3 and CHO cells are cultured with Dulbecco's Modified Eagle Medium and RPMI1640 with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin. HUVEC was cultured with Endothelial Cell Growth Medium (EGM-2). Aspirin (ASA), sialic acid (SA), 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay reagent, and dimethyl sulfoxide (DMSO) were procured from Sigma-Aldrich, whereas endotoxin-free recombinant human APE1/Ref-1 protein (rhAPE1/Ref-1; #MR-EAPE-100), monoclonal (clone 14E1), and polyclonal antibodies targeting APE1/Ref-1 were purchased from MediRedox. Additional antibodies were acquired from Cell Signaling Technology, Enzo Life Sciences, and Santa Cruz Biotechnology, and PD SpinTrap G-25 was obtained from GE Healthcare Life Sciences. ASA was dissolved in distilled water, and the pH was adjusted to 7.4 using sodium hydroxide; rhAPE1/Ref-1 was reconstituted in phosphate-buffered saline (PBS).

PEO-14 ovarian cancer cells were cultured in RPMI 1640 medium supplemented with 10% FBS, 1 mM sodium pyruvate, and 1% antibiotic-antimycotic solution. Additionally, PEO-14 cells were cultured as spheroids in a 1:1 mixture of MCDB 105 and Medium 199.

Purified recombinant human APE1/Ref-1 protein (100 µg) was subjected to in vitro acetylation by incubating it with 17.5 mM ASA solution for 4 h. Unreacted ASA was then removed using PD SpinTrap G-25 desalting spin columns (GE Healthcare Life Sciences), as described previously [10]. This process yielded approximately 90% acetylated APE1/Ref-1 protein, which was subsequently analyzed by immunoblotting and immunoprecipitation.

Recombinant receptor for advanced glycation end-product (RAGE) protein (R&D System) was incubated with either acetylated recombinant APE1/Ref-1 or recombinant APE1/Ref-1 (MediRedox) protein at a 1:1 ratio in binding buffer (20 mM Tris-HCl, 0.1% Triton X-100, 5% glycerol, 10 mM butyrate, 10 mM nicotinamide, 5 µM Trichostatin A) at 4°C for 1 h. Subsequently, pull-down with an anti-RAGE antibody (Santa Cruz Biotechnology) was performed by overnight incubation at 4°C. Bound proteins were followed using immune precipitation with SDS sample buffer and analyzed using Western blotting

Cell viability was determined using the MTT assay [21]. PEO-14 cells were seeded in 96-well plates and treated with rhAPE1/Ref-1 or ASA for 24 h. The MTT reagent (10 µl, 5 mg/ml) was added and incubated for 2 h at 37°C. Formazan crystals were dissolved in DMSO (100 µl), and absorbance was measured at 600 nm using a microplate reader (GloMax Discover, Promega).

Spheroid formation was initialed by seeding 5,000 cells into each well of a 96-well U-bottom plate with a non-adherent surface (Nunclon Sphara 96U-well plate #179425, Thermo Fisher Scientific). Cells were gently centrifuged at 400 × g for 5 min to promote spheroid formation. After 72 h of incubation to allow spheroid formation, the spheroids were treated with ASA (3 mM) and/or recombinant human APE1/Ref-1 (rhAPE1/Ref-1) (5 µg/ml) for an additional 24 h. Spheroid viability was assessed using both the Cyto3D Live-Dead Assay Kit (Well Bioscience, BM01) and the CellTiter-Glo 3D Cell Viability Assay (Promega) according to the manufacturer’s protocols. For the Cyto3D assay, 2 µl of reagent was added per 100 µl of total well volume, which included the culture medium. The spheroids were incubated with the reagent at 37°C for 10 min, and live (AO⁺, green) and dead (PI⁺, red) cells were visualized under fluorescence microscopy. For the CellTiter-Glo assay, luminescence corresponding to cellular ATP levels was measured using a luminometer, providing an additional quantitative assessment of spheroid viability.

Total RNA was extracted from PEO-14 cells using the RNeasy mini kit (Qiagen) following the manufacturer's instructions. The concentration and purity of RNA were determined using a NanoDrop spectrophotometer (Thermo Fisher Scientific). Complementary DNA was synthesized from 1 µg of total RNA using a Reverse Transcription PCR Kit (iNtRON Biotechnology). qRT-PCR was performed using SYBR Green Master Mix (Promega) and specific primers for the RAGE (forward: 5’- GTGTCCTTCCCAACGGCTC-3’, reverse: 5’-ATTGCCTGGCACCGGAAAA-3’) on a QuantStudio 5 Real-Time PCR System (Thermo Fisher Scientific). The thermal cycling conditions were as follows: an initial denaturation step at 95°C for 10 min, followed by 40 cycles of 95°C for 10 sec and 60°C for 30 sec. Relative mRNA expression levels were quantified using the ΔCt method, with β-actin (forward: 5’-CATGTACGTTGCTATCCAGGC-3’, reverse: 5’-CTCCTTAATGTCACGCACGAT-3’) as the internal control. All reactions were performed in triplicate to ensure reproducibility.

Flow cytometry using the annexin V-fluorescein isothiocyanate (FITC)/propidium iodide (PI) staining was used to monitor apoptosis [22]. Briefly, the cells were washed, trypsinized, and resuspended in the staining solution provided with the Annexin V-FITC Apoptosis Detection Kit (Abcam) following the manufacturer’s protocol. After incubation for 10 min, apoptosis was assessed using a flow cytometer (FACSCanto; BD Bioscience).

PEO-14 cells were seeded on a coverslip in 12-well plates. The Seeded cells were treated with 3 mM ASA for 6 h and then fixed with 4% freshly prepared paraformaldehyde in PBS for 15 min at room temperature. Fixed cells were washed with PBS and permeabilized with 0.1% Triton X-100 in PBS for 10 min. Following permeabilization (or no permeabilization), the cells were washed with bovine serum albumin (BSA) buffer (0.15% glycine and 0.5% BSA in PBS) and then blocked with 5% BSA buffer in PBS for 45 min at room temperature to minimize non-specific binding. Primary antibodies to RAGE (Santa Cruz Biotechnology) were diluted in 0.5% BSA buffer and incubated with the cells overnight at 4°C. The cells were washed with BSA buffer and then incubated with Alexa Fluor 488 (Thermo Fisher Scientific) secondary antibodies diluted in 0.5% BSA buffer for 1 h at room temperature in the dark. Nuclei were counterstained with 4',6-diamidino-2-phenylindole (DAPI) for 3 min. The coverslips were mounted onto glass slides using an antifade mounting medium. Fluorescent images were captured using an Axiophot fluorescence microscope (Carl Zeiss) with excitation/emission wavelengths of 488/520 nm.

PEO-14 cells (1 × 10⁵ cells/coverslip) were seeded on coverslips and treated with ASA and/or rhAEP1/Ref-1 to induce apoptosis. After 24 h, cells were stained with annexin V-FITC (5 µl) and PI (1 µg/ml), fixed in 2% formaldehyde, and counterstained with DAPI. Images were captured using fluorescence microscopy.

Caspase-3/7 activity was measured using the Caspase-Glo 3/7 assay kit (Promega) as per the instruction manual. Luminescence, indicating caspase-3/7 activity, was measured using a GloMax Luminometer (Promega).

Immunoprecipitation using an anti-acetyl lysine antibody was performed as previously reported [23], with some modifications, to analyze in vitro acetylation and protein binding. Immunoprecipitation using an anti-acetyl lysine antibody was performed to analyze the acetylated APE1/Ref-1 or RAGE binding. Briefly, 1 µg of each antibody was added and incubated for 2 h at 4°C. Protein A/G agarose beads were then added to each sample and incubated for 18 h at 4°C. The immunocomplex was collected using centrifugation at 300 rpm for 3 min and washed three times in a washing buffer. Each sample was subjected to 10%–12% SDS-PAGE, which was followed by immunoblotting using anti-APE1/Ref-1 antibody or anti-RAGE antibody.

ASA-treated or control PEO-14 cells were harvested with buffer A (250 mM sucrose, 1 mM EGTA, 10 mM HEPES, protease inhibitor, pH 7.5) and homogenized for 10 sec. Subsequently, samples were centrifuged at 3,000 rpm for 15 min to collect the supernatant. The supernatant was transferred to buffer B (1 M Na₂CO₃) and rotated for 45 min at 4°C. To re-collect the plasma membrane, the samplers were centrifuged at 12,000 rpm for 15 min to remove the supernatant. The pellet was then resuspended in buffer C (250 mM sucrose, 1 mM MgCl₂, and 10 mM HEPES).

Cells were washed with DPBS and lysed in RIPA buffer containing protease inhibitors. The lysates (30 µg) were separated into 10% SDS-PAGE gels. After transfer to polyvinylidene fluoride membranes and blocking with 5% non-fat dry milk in TBS-T, the membranes were probed overnight at 4°C with antibodies against acetyl-lysine (Cell signaling Technology, Cat. #9441), PAPR (Enzo Life Science, Cat. #SA-250), APE1/Ref-1 (MediRedox, Cat. #MR-PAAPE), Caspase-3 (Cell signaling Technology, Cat. #9664), RAGE (Santa Cruz Biotechnology, Cat. #sc-365154), N-cadherin (Abcam, Cat. #22744), and β-actin (Sigma-Aldrich, Cat. #A5316). Following incubation with HRP-conjugated secondary antibodies, chemiluminescence signals were detected using SuperSignal West Pico or Femto Substrate (Thermo Fisher Scientific) and protein levels were quantified by densitometry relative to β-actin. The plasma membrane of PEO-14 cells was prepared by sucrose density gradient centrifugation as described previously [10]. The membrane pellet was analyzed by immunoblotting using anti-RAGE and anti-N-cadherin antibodies (as plasma membrane loading control).

RAGE expression was knocked down using RNA interference (RNAi). Briefly, PEO-14 cells were transfected with either a control siRNA or a specific siRNA targeting RAGE using Lipofectamine RNAiMAX transfection reagent (Invitrogen) according to the manufacturer’s instructions. After 48 h, the knockdown efficiency was confirmed by Western blot analysis. The transfected cells were then subjected to the indicated treatments and assays as described above.

All statistical analyses were performed using GraphPad Prism software version 8 (GraphPad Software). The statistical significance of the differences was determined using a paired t-test or one-way analysis of variance, followed by Dunnett’s or Bonferroni multiple comparison tests. Differences were considered statistically significant at p < 0.05.

First, we investigated the effect of aspirin (ASA) on rhAPE1/Ref-1 acetylation and cellular hyperacetylation. rhAPE1/Ref-1 was treated with either ASA or SA (a negative control). Immunoprecipitation and Western blotting using an acetyl lysine antibody revealed that ASA treatment significantly increased rhAPE1/Ref-1 acetylation compared to SA treatment, indicating that rhAPE1/Ref-1 acetylation was ASA-specific (Fig. 1A). PEO-14 cells were treated with ASA and analyzed by Western blotting using an acetyl lysine antibody. The results showed markedly increased acetylation of various cellular proteins upon ASA treatment (Fig. 1B), confirming that ASA induces widespread cellular protein hyperacetylation. The stability of the acetyl group in acetylated rhAPE1/Ref-1 (Ac-rhAPE1/Ref-1) was assessed over time (0–24 h) using western blot analysis with acetyl-lysine and APE1/Ref-1 antibodies. The acetyl group was retained for up to 12 h, after which the acetyl-lysine signal significantly decreased, but the APE1/Ref-1 signal remained constant (Fig. 1C, D). In contrast, when rhAPE1/Ref-1 was co-incubated with ASA, acetylation was rapidly induced within 3 h and remained stable for the full 24 h observation period (Fig. 1E, F). This suggests that the acetyl modification is not permanent and may require co-administration of ASA for long-term maintenance of Ac-rhAPE1/Ref-1.

Given that ASA induced cellular hyperacetylation and the in vitro acetylation of rhAPE1/Ref-1, we investigated the combined effects of rhAPE1/Ref-1 or Ac-rhAPE1/Ref-1 and aspirin in ovarian cancer cells. PEO-14 cells were treated with either protein (0–5 µg/ml) and/or ASA (3 mM) for 24 h. Cell viability was then evaluated using the MTT assay. As shown in Fig. 2A, B, neither rhAPE1/Ref-1 nor Ac-rhAPE1/Ref-1 alone induced significant changes in the viability of PEO-14 cells. However, in PEO-14 cells pre-exposed to ASA for 6 h, Ac-rhAPE1/Ref-1 treatment led to reduced cell viability at high concentrations (5 µg/ml) (Fig. 2C). Notably, co-treatment of PEO-14 cells with rhAPE1/Ref-1 and ASA reduced cell viability. Although ASA (3 mM) alone did not affect cell viability (Fig. 2D), co-treatment with rhAPE1/Ref-1 (≥ 0.5 µg/ml) and ASA resulted in a dose-dependent increase in PEO-14 cell death (Fig. 2D). Moreover, the combination of 5 µg/ml rhAPE1/Ref-1 and 3 mM ASA induced approximately 40% cell death in PEO-14 cells. Furthermore, the effect of the combination of ASA and rhAPE1/Ref-1 on cell viability in other HGSOC cell lines, CAOV3, and non-cancerous cell lines, including CHO and HUVEC cells. In those cell lines, no significant reduction in cell viability was observed with either ASA alone, rhAPE1/Ref-1 alone, or the combination of ASA and rhAPE1/Ref-1 (0–5 µg/ml) (Supplementary Fig. 1). This result suggests that ASA-induced cellular hyperacetylation may create a cellular environment where APE1/Ref-1, particularly in its acetylated form, reduces cell viability in PEO-14 ovarian cancer cells.

PEO-14 cells were treated with ASA (3 mM), and RAGE expression was assessed by Western blotting at various time points (0–24 h) to investigate whether ASA affects RAGE expression. ASA treatment in PEO-14 cells leads to a time-dependent increase in RAGE protein expression, with a significant increase observed as early as 3 h after treatment and peaking at 12 h (Fig. 3A, B). To determine whether ASA-induced RAGE expression also occurs at the transcriptional level, RAGE mRNA expression was analyzed by real-time quantitative RT-PCR (qRT-PCR). The results showed that there was no change in RAGE mRNA expression within the first 6 h of ASA treatment; however, RAGE mRNA expression began to increase after 12 h of treatment (Fig. 3C). In contrast, RAGE expression did not significantly increase in other cell lines, including CAOV3, CHO, and HUVEC, following ASA treatment (Supplementary Fig. 2), suggests that the cellular response to ASA-induced RAGE expression may vary across different cell types.

To investigate whether ASA affects the intracellular translocation of RAGE in PEO-14 cells, immunofluorescence staining was performed on both permeabilized and non-permeabilized cells (Fig. 3D) [24]. In permeabilized cells, RAGE was primarily expressed in the nucleus. Treatment with ASA (3 mM, 6 h) induced the translocation of RAGE from the nucleus to the cytoplasm. In non-permeabilized cells, RAGE expression was not detected under control conditions, but it was observed after ASA treatment (3 mM, 6 h). These results suggest that ASA promotes the cell surface expression of RAGE. Western blot analysis of plasma membrane fractions revealed that ASA treatment increased RAGE in the plasma membrane, with N-cadherin serving as a control for membrane protein enrichment (Fig. 3E). Quantification of RAGE band intensity in the plasma membrane fraction from Western blot analysis is shown in Fig. 3F. These findings suggest that ASA not only induces the expression of RAGE but also promotes its translocation to the cell surface, indicating a potential role for ASA in modulating RAGE signaling pathways.

Co-immunoprecipitation assays were performed to investigate the interaction between rhAPE1/Ref-1 and RAGE. Briefly, 1 µg of rhRAGE was incubated with either rhAPE1/Ref-1 or Ac-rhAPE1/Ref-1 for 6 h. The resulting complexes were immunoprecipitated using an anti-RAGE antibody and analyzed by western blotting. As shown in Fig. 4A, rhAPE1/Ref-1 did not co-precipitate with RAGE, indicating no interaction. In contrast, a clear band corresponding to Ac-rhAPE1/Ref-1 (39 kDa) was detected in the RAGE immunoprecipitate, demonstrating a strong and specific interaction between Ac-rhAPE1/Ref-1 and RAGE. This result highlights the importance of acetylation in mediating the binding between APE1/Ref-1 and RAGE, and strongly suggests that RAGE is a putative receptor for acetylated APE1/Ref-1. Additionally, the ASA-induced increase in RAGE expression was effectively inhibited by RAGE siRNA treatment (Fig. 4B). Subsequently, we examined the effect of rhAPE1/Ref-1 and ASA co-treatment in cells where RAGE expression was knocked down by siRNA to investigate the role of RAGE signaling in the reduced cell viability of PEO-14 ovarian cancer cells caused by this co-treatment. As shown in Fig. 4C, RAGE knockdown significantly inhibited the reduction in cell viability induced by co-treatment of rhAPE1/Ref-1 with ASA. These results indicate that ASA-induced RAGE expression is crucial for the reduced cell viability caused by the combination of APE1/Ref-1 with ASA. To investigate the role of RAGE in acetylated APE1/Ref-1-induced caspase activation, we knocked down RAGE expression in PEO-14 cells using siRNA. Western blot analysis revealed that RAGE knockdown significantly attenuated acetylated APE1/Ref-1-induced cleavage of PARP or caspase-3, indicating a critical role of RAGE in mediating PARP activation downstream of acetylated APE1/Ref-1 (Fig. 4D, E).

We examined whether rhAPE1/Ref-1 and ASA trigger apoptosis, caspase activation, and PARP cleavage in PEO-14 ovarian cancer cells to investigate the mechanism underlying their synergistic effect in inducing ovarian cancer cell death. Immunofluorescence imaging of PEO-14 cells following co-treatment with rhAPE1/Ref-1 and ASA revealed a marked increase in the abundances of both annexin V–positive cells and PI-stained cells compared to that seen with the abundances of untreated cells or cells treated with ASA alone (Fig. 5A). This observation suggests that co-treatment of rhAPE1/Ref-1 with ASA induces apoptosis in PEO-14 cells.

Flow cytometry analysis further corroborated this finding, demonstrating a concentration-dependent increase in the percentage of total apoptotic cells (both early and late apoptotic) upon treatment with increasing concentrations of rhAPE1/Ref-1 in the presence of 3 mM ASA (Fig. 5B, C). Notably, co-treatment with 5 µg/ml rhAPE1/Ref-1 and 3 mM ASA induced significant apoptosis in approximately 33.6% of PEO-14 cells. In contrast, treatment with ASA alone did not significantly alter the apoptosis rate compared to that in the untreated control, indicating that the observed pro-apoptotic effect is specific to the combined treatment.

We then examined the activation of caspase-3/7, key caspases involved in apoptosis, to investigate the molecular mechanism underlying the observed apoptosis. Although ASA alone did not affect caspase-3/7 activity, co-treatment with rhAPE1/Ref-1 and ASA significantly increased caspase-3/7 activity in a dose-dependent manner (Fig. 5D). In addition, we assessed the cleavage of PARP, a DNA repair enzyme and a well-known substrate of activated caspase-3. Western blot analysis revealed a notable increase in the cleavage of both PARP and caspase-3 in PEO-14 cells co-treated with rhAPE1/Ref-1 and ASA, whereas ASA alone had no effect (Fig. 5E). These results confirm that the combined treatment activates the caspase cascade, leading to PARP cleavage and ultimately resulting in apoptotic cell death.

We employed 3D spheroid cultures of PEO-14 ovarian cancer cells to evaluate the effect of the combined treatment in a more physiologically relevant setting. As shown in Fig. 6A, treatment with either ASA (3 mM) or rhAPE1/Ref-1 (5 µg/ml) alone did not significantly induce cell death in PEO-14 spheroids, as evidenced by minimal PI staining. However, co-treatment with rhAPE1/Ref-1 and ASA led to a marked increase in PI-positive cells, indicative of apoptosis (Fig. 6B). We further assessed the impact on spheroid viability via CellTiter-Glo 3D Cell Viability Assay. Consistent with the immunofluorescence data, co-treatment with rhAPE1/Ref-1 and ASA resulted in a significant decrease in spheroid viability compared to that seen with either treatment alone (Fig. 6C).

These findings demonstrate that the combination of rhAPE1/Ref-1 and aspirin effectively induces cell apoptosis in 3D spheroid cultures of PEO-14 ovarian cancer cells, supporting the therapeutic potential of acetylated APE1/Ref-1 as a novel therapeutic strategy.